The authors declare no competing financial interests.

All animal procedures were approved by the Animal Welfare Department of the Th&#252;ringer Landesamt f&#252;r Lebensmittelsicherheit und Verbraucherschutz (03-048/16; TLV; Bad Langensalza, Germany).
1. Cardiotoxin-induced Muscle Injury
NOTE: Perform all experiments involving living mice in accordance with the national Animal Welfare Act and under appropriate aseptic conditions. Also, sacrificing of animals must be performed in accordance with the national Animal Welfare Act.
<ol>
	<li>Disinfect the inhalation box used for inhalation anesthesia and the surgery area with 70% ethanol. Place a sterile surgical cloth on the heating pad where the surgery will take place.</li>
	<li>Turn on the heating pad at 37 &#176;C.</li>
	<li>Administer analgesics (e.g., 1 mg/kg body weight meloxicam subcutaneously) 15 min prior to start of the surgery. 
	NOTE: For a typical experiment, use mice which are at least 6 weeks old, sex-matched and in a good general physical condition.</li>
	<li>Prepare the cardiotoxin solution (20 &#181;M in 0.9 % NaCl), store the solution at -20 &#176;C.</li>
	<li>Allow the cardiotoxin solution to reach room temperature (RT).</li>
	<li>Transfer the mouse into the inhalation box and induce anesthesia with isoflurane (initiation 3.5 &#8211; 5 % in pure oxygen). Check the depth of anesthesia with a lack of response to a toe pinch.</li>
	<li>Place the mouse on a clean paper towel and maintain anesthesia with a nose mask (1.5 &#8211; 3 % isoflurane in pure oxygen). Shave the injection area of the cranial lower leg (from knee to paw, Figure 1A). Remove all excessive loose hair. 
	NOTE: Physically separating the shaving and injection area in the room will provide more sterile conditions for the injections.</li>
	<li>Place the mouse in the lateral recumbent&#160;position on the heating pad covered with a sterile surgical cloth and maintain anesthesia with a nose mask (1.5 &#8211; 3% isoflurane in pure oxygen).</li>
	<li>Disinfect the injection area of the cranial lower leg (from knee to paw) with 70% ethanol.</li>
	<li>Perform a toe pinch before starting the intramuscular injection to ensure an adequate depth of anesthesia.</li>
	<li>Inject 50 &#181;L cardiotoxin (20 &#181;M in 0.9% NaCl) into the tibialis anterior muscle using an insulin syringe with a 29-gauge needle. First, pierce the skin just distal of the knee.</li>
	<li>Insert the needle fully into the muscle (in myofiber orientation/parallel to the tibia bone) and inject the cardiotoxin slowly (10-20 s) along the full length of the muscle while moving the needle back and forth to allow an even distribution of the cardiotoxin thereby injuring the whole tibialis anterior muscle (Figure 1B,C). 
	NOTE: Injure the tibialis anterior muscle from only one leg, the contralateral tibialis anterior muscle can serve as an internal control to determine that skeletal muscles were not pathologically affected before cardiotoxin injury.</li>
	<li>Transfer the mouse back to its cage placed on a heating pad and monitor the recovery process until the animal is conscious and becomes ambulatory. 
	NOTE: The mice will only show a slight limp. If mice are limping heavily and do not put weight on the leg at all, sacrifice the mouse.</li>
	<li>Administer analgesics during the following 2 days (e.g., 1 mg/kg body weight meloxicam subcutaneously, every 24 h) and monitor them on a weekly basis.</li>
</ol>
2. Injection of Self-delivering siRNA into the Regenerating Tibialis Anterior Muscle (At Day 3 Post Cardiotoxin Injury)
<ol>
	<li>Prepare the siRNA solution by resuspending the siRNA (e.g., siRNA smart pool) in 0.9% NaCl (final concentration of 2 &#181;g/&#181;L) according to the manufacturer&#8217;s instruction. 
	NOTE: The siRNA is chemically modified to facilitate the passive uptake by cells and protect it from nuclease degradation. No transfection reagents are needed, thereby reducing toxicity. Using a smart pool of four independent siRNA sequences targeting the same gene increases knockdown efficiency.</li>
	<li>Store the siRNA at -20 &#176;C and transfer it on ice to the surgery room.</li>
	<li>Disinfect the inhalation box used for inhalation anesthesia and the surgery area with 70% ethanol. Place a sterile surgical cloth on the heating pad where the surgery will take place.</li>
	<li>Turn on the heating pad at 37 &#176;C.</li>
	<li>Transfer the mouse into the inhalation box and induce anesthesia with isoflurane (initiation 3.5 &#8211; 5% in pure oxygen). Check the depth of anesthesia with a lack of response to a toe pinch.</li>
	<li>Place the mouse in the lateral recumbent&#160;position on the heating pad covered with a sterile surgical cloth and maintain anesthesia with a nose mask (1.5 &#8211; 3% isoflurane in pure oxygen).</li>
	<li>Disinfect the injection area of the cranial lower leg (from knee to paw).</li>
	<li>Perform a toe pinch before starting the intramuscular injection to ensure an adequate depth of anesthesia.</li>
	<li>Inject up to 50 &#181;L siRNA solution (up to 100 &#181;g total in 0.9% NaCl; directed against the target gene; use scrambled siRNA as control) into the injured tibialis anterior muscle using an insulin syringe with a 29 G needle. First, pierce the skin just distal of the knee, then insert the needle into the tibialis anterior muscle.</li>
	<li>Insert the needle fully into the muscle (in myofiber orientation/parallel to the tibia bone) and inject the siRNA solution slowly (10 &#8211; 20 s) along the full length of the muscle while moving the needle back and forth to allow an even distribution of the siRNA along the whole tibialis anterior muscle.</li>
	<li>Transfer the mouse back to its cage placed on a heating pad and monitor the recovery process until the animal is conscious and becomes ambulatory. 
	NOTE: Analgesia is not necessarily following the injection of siRNA.</li>
</ol>
3. Dissection of Tibialis Anterior Muscle
<ol>
	<li>Prepare the freezing solution (2 parts OCT compound and 1 part 30% sucrose in deionized water) at least 12 h before usage to avoid air bubbles.</li>
	<li>Prepare freezing molds by wrapping an aluminum foil around a pencil and seal it with tape. It is important that the bottom of the mold provides an even, closed surface. 
	NOTE: Smaller freezing molds allow the faster freezing of the muscle avoiding freezing artifacts.</li>
	<li>Sacrifice the mouse, e.g., by CO2 inhalation, at the respective time point of regeneration and spray the whole mouse and dissection tools with 70% ethanol. 
	NOTE: Cervical dislocation can be applied, in addition, to avoid excessive bleeding at the leg when isolating the tibialis anterior muscle.</li>
	<li>Remove the fur and skin from the injured hindlimb by cutting the skin at the ankle with extra fine sharp scissors (cutting edge: 13&#160;mm) and pulling up the skin towards the knee using forceps.</li>
	<li>To expose the tibialis anterior tendon at the ankle, pull the remaining skin towards the foot or cut with sharp scissors. 
	NOTE: The mouse can be pinned to a support board to allow better fixation.</li>
	<li>Before harvesting the tibialis anterior muscle, remove the fascia using fine forceps (Dumont 5 or 7, straight or curved). Pinch the closed forceps through the fascia next to the tibia bone at the ankle of the injured leg (Figure 2A). Move the forceps towards the knee thereby tearing the fascia and exposing the tibialis anterior muscle (Figure 2B).</li>
	<li>To isolate the tibialis anterior muscle, expose the distal tendon and grab it with fine forceps (Dumont 7, curved). Cut the tendon using spring scissors (cutting edge: 5&#160;mm, tip diameter: 0.35&#160;mm) and pull the muscle (holding it at the tendon) towards the knee.</li>
	<li>To harvest the tibialis anterior muscle, cut it as close to the knee as possible using sharp scissors.</li>
	<li>Prior to freezing, cut the tibialis anterior muscle at the mid belly region of the muscle with straight scissors into two halves of equal size to allow the analysis of the mid belly region (Figure 2C,D).</li>
	<li>Fill the freezing mold halfway with the freezing solution.</li>
	<li>Insert the two halves of the tibialis anterior muscle into the freezing mold with the mid belly region facing the bottom of the mold (Figure 2E). Make sure the tibialis anterior halves are inserted in an upright position leaning against the wall of the freezing mold to avoid tipping of the muscle. 
	NOTE: The more freezing solution is in the mold, the longer the freezing will take, thus increasing the risk of cryo-artifacts.</li>
	<li>Hold the freezing mold using forceps and transfer it halfway into liquid nitrogen (Figure 2F). Make sure that no liquid nitrogen is entering the freezing mold during the freezing process.</li>
	<li>Observe the freezing process, the freezing medium changes color from transparent to white and becomes solid. Submerse the freezing mold in liquid nitrogen for a few seconds and transfer the freezing mold to a -80 &#176;C freezer or to dry ice for future processing.</li>
	<li>Store the frozen muscles in the freezing molds at -80 &#176;C until further usage. 
	NOTE: The freezing molds fit well in 24-well plates or 1.5 mL reaction tubes allowing labeling and organized storage.</li>
	<li>Handle the muscle in the freezing molds for further usage always on dry ice.</li>
</ol>
4. Cryosectioning of Regenerating Tibialis Anterior Muscle
<ol>
	<li>Before starting sectioning, set the chamber temperature of the cryostat to -21 &#176;C, the object temperature to -20 &#176;C and the cutting thickness to 10, 12 or 14 &#181;m (depending on the future applications) (Figure 3A).</li>
	<li>Transfer the muscle sample in the freezing mold into the cryostat and let it adjust to the temperature for several minutes. During this time, label the microscope slides.</li>
	<li>Remove the foil around the embedded muscle using precooled forceps inside the cryostat. Avoid touching of the sample as the cryomedium starts to thaw easily.</li>
	<li>Mount the sample with the mid belly region of the tibialis anterior muscle directed upwards onto the metal sample holder of the cryostat using cryomedium. This ensures that the cutting site of the muscle (bottom of the mold) is facing the experimenter.</li>
	<li>Insert the sample holder with the mounted sample into the sectioning mechanism. 
	NOTE: To ensure proper sectioning of the sample, use a new blade before starting.</li>
	<li>First, trim the sample block (30 &#181;m) to get an even sectioning plane and to reach the respective region of the muscle (Figure 3B).</li>
	<li>After trimming, cut consecutive sections of the respective thickness (e.g., 14 &#181;m).</li>
	<li>Collect the sections on microscope glass slides by holding the slide face down over the section. The section is attaching to the slide (Figure 3C).</li>
	<li>Store sections at -80 &#176;C or -20 &#176;C until further use or directly use for immunofluorescence or immunohistochemistry.</li>
</ol>
5. Immunostaining for Markers of Regeneration
<ol>
	<li>Perform all further steps in a humidified chamber.</li>
	<li>Briefly, equilibrate the sections at room temperature.</li>
	<li>Fix sections with 1 mL of 2% PFA (in PBS, pH 7.4) per slide for 5 min at RT.</li>
	<li>Remove the 2% PFA solution by pouring it into the respective waste container.</li>
	<li>Wash 3 times with 1 mL of PBS (pH 7.4) for 5 min at RT.</li>
	<li>Remove the PBS by pouring it into a waste container.</li>
	<li>Per slide, add 1 mL of the permeabilization solution (0.1 % Triton X-100, 0.1 M Glycine in PBS pH 7.4) for 10 min.</li>
	<li>Remove the permeabilization solution by pouring it into a waste container.</li>
	<li>Add 150 &#181;L of the blocking solution (M.O.M. 1:40 in PBS pH 7.4) per slide and cover with the coverslip. Incubate for 1 h at RT.</li>
	<li>Remove the coverslip and the blocking solution, apply 100 &#181;L of primary antibody solution [PAX7, DSHB, mouse IgG1, undiluted or devMHC, DSHB, undiluted, mouse IgG1and laminin (rabbit, 1:1,000)] per slide. Cover with coverslip. Incubate O/N (overnight) at 4 &#176;C. 
	NOTE: Perform a control staining by omitting the primary antibodies, incubate the section with blocking solution instead.</li>
	<li>Wash 3 times with 1 mL of PBS (pH 7.4) for 5 min at RT.</li>
	<li>Remove PBS by pouring it into a waste container.</li>
	<li>Add 100 &#181;L of the secondary antibody solution (Alexa Fluor 546 goat anti-mouse IgG1 and Alexa Fluor 488 donkey anti-rabbit IgG in blocking solution, 1: 1,000) per slide and cover with the coverslip. Incubate for 1 h at RT.</li>
	<li>Incubate the slides in the dark, e.g., use an aluminum foil to cover or use a black humidified chamber. 
	NOTE: From now on all steps should be performed in reduced light conditions since some secondary antibodies are light sensitive.</li>
	<li>Remove the coverslip and the secondary antibody solution.</li>
	<li>Wash 3 times with 1 mL of PBS (pH 7.4) for 5 min at RT.</li>
	<li>Remove the PBS by pouring it into a waste container.</li>
	<li>Perform DAPI staining by adding 1 mL of the solution per slide to a final concentration of 10 &#181;g/mL for 5 min at RT.</li>
	<li>Remove the DAPI staining solution by pouring it into a waste container.</li>
	<li>Wash 3 times with 1 mL PBS (pH 7.4) for 5 min at RT.</li>
	<li>Completely remove the PBS used for washing.</li>
	<li>Apply 2 &#8211; 3 drops of aqueous mounting medium and directly cover the slide with a new coverslip.</li>
	<li>Let the slides dry for 1 h at RT in the dark.</li>
	<li>Store the stained sections at 4 &#176;C in the dark until further analysis using a fluorescence microscope.</li>
</ol>
6. Hematoxylin and Eosin Staining
<ol>
	<li>Perform all further steps in a humidified chamber.</li>
	<li>Fix sections with 1 mL of 2% PFA (in PBS, pH 7.4) per slide for 5 min at RT.</li>
	<li>Remove the 2% PFA solution by pouring it into the respective waste container.</li>
	<li>Transfer the slides into a Coplin jar.</li>
	<li>Gently rinse the slides with tap water. 
	NOTE: Do not turn on the faucet too high since that might damage the sections.</li>
	<li>Transfer the slides to the humidified chamber. Remove the liquid.</li>
	<li>Apply 1 mL of Hematoxylin staining solution (Gill No 3) for 2 min.</li>
	<li>Transfer the slides into a Coplin jar.</li>
	<li>Gently rinse the slides with the tap water until the nuclei turn blue. 
	NOTE: Do not turn on the faucet too high since that might damage your sections.</li>
	<li>Transfer the slides to the humidified chamber. Remove the liquid.</li>
	<li>Apply 1 mL of Eosin Y solution and incubate for 2 min.</li>
	<li>Transfer the slides into a Coplin jar.</li>
	<li>Gently rinse the slides with tap water until the tap water coming off the slides is clear.</li>
	<li>Remove all liquid and let the slides dry for 2 min.</li>
	<li>Mount in mounting medium suitable for immunohistochemistry.</li>
</ol>

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We would like to thank Christina Picker and Christine Poser for excellent technical assistance and Saskia Steiner for help with the determination of siRNA transfection efficiency. We thank the core service histology for great technical support, especially Sabine Landmann and Linda Rothenburger. We also thank the animal facility mouse at the FLI for excellent support. This work was supported by grants from the Deutsche Forschungsgemeinschaft (MA-3975/2-1) and the Deutsche Krebshilfe (DKH-JvM-861005) to J.v.M.

Here we present a method to investigate the function of a specific gene during regeneration of skeletal muscle without the need of transgenic animals. This is accomplished by the combination of cardiotoxin induced muscle injury with the injection of a self-delivering siRNA into the regenerating skeletal muscle at day 3 after injury. We have described in detail the procedures of muscle injury by cardiotoxin, the injection of the self-delivering siRNA and processing of the harvested muscles to analyze the progress of regeneration. We demonstrate that injection of the snake venom cardiotoxin into skeletal muscle effectively injures the whole muscle and that self-delivering siRNAs are found in around 75% of all satellite cells among other cell types two days after their injection into the regenerating skeletal muscle (Figure 4, Figure 5).
Particular attention should be focused on a homogeneous injury of the tibialis anterior muscle since varying degrees of injury influence the regeneration outcome and thereby also the effect of the siRNA might be affected. Furthermore, it is critically important that the whole regenerating area is injected with the self-delivering siRNA. For the analysis of the regeneration process, it is recommended to always compare similar areas of the regenerating skeletal muscle, therefore, the tibialis anterior muscle should be cut in half to always compare the mid-belly region of the muscle. When analyzing the muscle, the whole cryosection needs to be analyzed since myofiber composition differs in the tibialis anterior muscle and might therefore regenerate differently.
The function of satellite cells can be investigated by various experimental procedures including their culture on the adjacent isolated single myofibers17, by transplantation and by analyzing regeneration of skeletal muscle after induced injury18,19. Investigating the function of satellite cells by using an in vivo induced injury model, e.g., injection of cardiotoxin, provides the ability to analyze satellite cell function also in terms of their interaction with other cell types such as macrophages and investigation of the influence of systemic factors2. Injury of skeletal muscle can be accomplished by various means, e.g., eccentric exercise, freeze injury, injection of BaCl2 or injection of snake venoms such as cardiotoxin or notexin18. While eccentric exercise is probably the most physiological injury method, the injury to one particular muscle is only limited20. Freeze injuries can be applied when the migration of satellite cells towards the site of injury is the aim of the study or only a specific part of the muscle should be injured. Experimentally the disadvantage of freeze injuries is the open surgery which needs to be performed to apply the precooled metal probe. Injection of BaCl2 or snake venoms is the most dramatic method of injury, thereby challenging satellite cell function the most. Furthermore, the injection is minimally invasive, surgery time, in general, is less than five minutes and does not involve suturing, etc. thereby minimizing the risk of infections.
Muscle injury is mostly used to investigate the functional consequences of loss of gene functions, e.g., loss of Pax77,21. Especially if aged mice are the focus of the scientific question, the generation or use of transgenic mice is often not feasible. Injection of self-delivering siRNAs targeting a specific gene is a viable alternative in those cases and has been used successfully22. In brief, the tibialis anterior muscle of mice was injured by injection of cardiotoxin and self-delivering siRNAs directed against fibronectin (FN) were injected at day 3 after injury. Muscles were analyzed 10 days after injury and a significant decrease in satellite cell numbers was observed in siFN versus scrambled siRNA control conditions. The knockdown efficiency was determined in whole muscle lysates by quantitative Real Time PCR 2 days after siRNA injection, a reduction in expression levels of 58% was achieved suggesting that delivery and knockdown efficiency are sufficient for functional analyses22. Alternatives for testing the knockdown efficiency are either immunoblot or immunofluorescence analyses with antibodies directed against the target gene. The efficiency and specificity of the siRNA used for in vivo injections should be determined before injection into mice, e.g., by testing the efficiency in isolated satellite cells or primary myoblasts. The use of a smart pool consisting of 4 different siRNAs versus a single siRNA increases the efficiency of knockdown but also increases the risk of unspecific targeting. The specificity of all siRNA sequences used should be tested in cell culture to avoid off-target effects. As a control, a non-targeting scrambled siRNA should be used since the injection of a siRNA per se might influence the regeneration process due to the injection and, thereby, additional damage of the muscle. The time point of injecting the siRNA is depending on the scientific question and on the expression profile of the target gene. Generally, an injection of self-delivering siRNA at day 3 after cardiotoxin injury targets most genes important for satellite cell proliferation since satellite cell proliferation peaks around day 3 after injury. The time point for the first siRNA injection should not be less than 48 h after cardiotoxin injury since the injection volume of cardiotoxin is quite high and reabsorption of the liquid should have taken place before injecting additional solutions into the muscle. In general, multiple injections of siRNAs or a combination of different siRNAs is possible although one must consider that each injection into the regenerating muscle is causing additional damage.
One limitation of the described method is the fact, that the effect observed is not necessarily only depending on knockdown of the target gene in satellite cells but might be attributed to other cell types such as immune cells or fibro-adipogenic progenitor cells. Therefore, it is necessary to combine those experiments with experiments investigating a pure population of satellite cells. One can either perform experiments using floating myofiber cultures, where satellite cells are cultured on their adjacent myofibers or perform transplantation experiments using isolated satellite cells17.
An alternative to the injection of self-delivering siRNAs is the injection of small molecule inhibitors or recombinant proteins which can be performed, depending on the scientific question. For instance, the injection of the extracellular matrix protein fibronectin or small molecule inhibitors of JAK/STAT signaling have been successfully performed in aged mice15,16. The analysis of a particular gene function in one specific cell type during regeneration of skeletal muscle, e.g., in satellite cells, is only possible through the use of an inducible genetic mouse model. Injections of self-delivering siRNAs, recombinant proteins or small molecule inhibitors might affect multiple cell types in regenerating skeletal muscle.

Skeletal muscle is the body&#8217;s largest tissue representing approximately 40% of total body weight enabling voluntary locomotion. The tissue architecture of skeletal muscle is mainly composed of postmitotic, terminally differentiated, multinucleated myofibers as well as various other cells from the peripheral nervous system, vascular system, and interstitial cells1. Importantly, skeletal muscle has a tremendous capacity to regenerate and restore function upon injury or damage2. This process depends on the tissue resident muscle stem cells also called satellite cells3,4. Satellite cells are located between the myofiber and the basal lamina and characterized by the expression of the transcription factor Pax75,6,7. Under homeostatic conditions, satellite cells are quiescent but get activated due to traumatic injury, e.g., through eccentric exercise or experimentally through injection of the snake venom cardiotoxin6,8. Once activated, Pax7 positive satellite cells co-express MyoD and Myf5, which commits the stem cells to myogenic differentiation. Satellite cells that resist the upregulation of commitment factors will retain their stemness potential and return to quiescence to replenish the stem cell pool for future demands. After the expansion of the myogenic progenitor pool, transcriptional networks are activated by differentiation factors like Myogenin to initiate the cell cycle exit and terminal differentiation. These myoprogenitors then fuse to each other or to existing myofibers contributing myonuclei to maintain the size of the myonuclear domain. The myofibers express terminal muscle differentiation genes like Myosin Heavy Chain. Finally, the newly formed myofibers grow and mature to build the functional units of skeletal muscle9,10.
The regeneration of skeletal muscle can be affected by various conditions including muscle diseases or aging11,12, ranging from mild impairments to life threatening conditions, e.g., in Duchenne muscular dystrophy13,14. Therefore, regenerative medicine aims to restore damaged or malfunctioning skeletal muscle tissue using its inherent regenerative power by targeting satellite cell function15,16. To use its full potential a comprehensive understanding of satellite cells in their endogenous niche during regeneration of skeletal muscle is required. Although experimental approaches exist to isolate satellite cells adjacent to their myofibers17, the full complexity of cellular and systemic interactions of satellite cells with their environment can only be recapitulated in vivo. In that regard, great knowledge about skeletal muscle regeneration has been acquired using mouse injury models2,18.
Here we introduce a specific experimental mouse injury model to study the stem cell mediated regeneration of cardiotoxin-induced damage of the tibialis anterior muscle in vivo. Cardiotoxin, a snake derived cytolytic toxin that causes myofiber depolarization and necrosis, is injected into the tibialis anterior muscle, which in turn will trigger tissue degeneration followed by regeneration. To analyze the function of genes during acute regeneration, self-delivering siRNAs are injected at day 3 after injury, at the peak of the satellite cell expansion. Experimental animals are sacrificed at various timepoints and the tibialis anterior muscles are collected. The dissected muscles are frozen and processed for further cryo-sectioning. Immunofluorescence microscopy is then used to analyze markers of regeneration. This method allows for the investigation of the function of a single gene during satellite cell driven regeneration of skeletal muscle using wild type mice.

<table><tbody><tr><td>Isoflurane</td><td>Henry Schein</td><td>Isothesia</td><td>inhalation narcotics</td></tr><tr><td>Hot Plate 062</td><td>Labotect</td><td>13854</td><td></td></tr><tr><td>Anesthesia System (Tec 7) with inhalation box + nose masks</td><td>Tem Sega</td><td>Minihub</td><td></td></tr><tr><td>Shaver for rodents</td><td>isis</td><td>GT420</td><td></td></tr><tr><td>Cardiotoxin</td><td>Latoxan</td><td>L8102</td><td>snake venom needed for muscle injury</td></tr><tr><td>Accell siRNA smart pool</td><td>Dharmacon</td><td>depending on your target gene</td><td>self delivering siRNA</td></tr><tr><td>Alexa Fluor 488 donkey anti-rabbit IgG</td><td>Thermo Fisher</td><td>A-21206</td><td>secondary antibody</td></tr><tr><td>Alexa Fluor 546 goat anti-mouse IgG1</td><td>Thermo Fisher</td><td>A-21123</td><td>secondary antibody</td></tr><tr><td>Coverslips</td><td>VWR</td><td>631-1574</td><td></td></tr><tr><td>CV Mount</td><td>Leica</td><td>14046430011</td><td>mounting medium for immunohistochemistry</td></tr><tr><td>DAPI</td><td>Sigma Aldrich</td><td>D9542</td><td>nuclear staining</td></tr><tr><td>devMHC antibody</td><td>DHSB</td><td>F.1652</td><td></td></tr><tr><td>Eosin Y</td><td>Thermo Fisher</td><td>73104</td><td></td></tr><tr><td>Haematoxylin Gill No3</td><td>Sigma-Aldrich</td><td>GHS316-500ML</td><td></td></tr><tr><td>Insulin syringe (29 g)</td><td>Terumo</td><td>3SS05M2813</td><td>syringue used for muscle injections</td></tr><tr><td>Laminin antibody</td><td>Sigma Aldrich</td><td>L9393</td><td></td></tr><tr><td>M.O.M blocking reagent</td><td>Vector labs</td><td>MKB-2213</td><td>blocking for immunofluorescent staining</td></tr><tr><td>Meloxicam</td><td>Boehringer Ingelheim</td><td>Metacam</td><td>analgesics</td></tr><tr><td>OCT</td><td>Thermo Fisher</td><td>6502</td><td>tissue embedding</td></tr><tr><td>Pax7 antibody</td><td>DSHB</td><td>PAX7</td><td>satellite cell specific antibody</td></tr><tr><td>ProLong Gold Antifade Mountant</td><td>Thermo Fisher</td><td>P36934</td><td>aequos mounting medium</td></tr><tr><td>Sucrose</td><td>Carl Roth</td><td>4621.1</td><td>tissue embedding</td></tr><tr><td>Superfrost plus</td><td>Thermo Scientific</td><td>J1830AMNZ</td><td>microscope slides</td></tr><tr><td>TritonX-100</td><td>Amresco</td><td>0694-1L</td><td>permeabilization reagent</td></tr><tr><td></td><td></td><td></td><td></td></tr><tr><td>&lt;u&gt;Dissection tools&lt;/u&gt;</td><td></td><td></td><td></td></tr><tr><td>Dumont 5, straight</td><td>Fine Science Tools</td><td>11295-10</td><td></td></tr><tr><td>Dumont 7, curved</td><td>Fine Science Tools</td><td>11272-40</td><td></td></tr><tr><td>Extra fine Bonn scissors (cutting edge: 13&amp;nbsp;mm)</td><td>Fine Science Tools</td><td>14084-08</td><td></td></tr><tr><td>Narrow pattern forceps</td><td>Fine Science Tools</td><td>11002-16</td><td></td></tr><tr><td>Spring scissors (cutting edge: 5&amp;nbsp;mm, tip diameter: 0.35&amp;nbsp;mm)</td><td>Fine Science Tools</td><td>91500-09</td><td></td></tr></tbody></table>

analyzing satellite cell function during skeletal muscle regeneration by cardiotoxin injury and injection of self-delivering sirna in vivo

Skeletal muscle possesses an enormous capacity to regenerate after injury. This process is mainly driven by muscle stem cells, also termed satellite cells. Satellite cells are characterized by the expression of the transcription factor Pax7 and their location underneath the basal lamina in the resting skeletal muscle. Upon injury, satellite cells get activated, undergo self-renewal or differentiation to either form new myofibers or to fuse with damaged ones. The functionality of satellite cells in vivo can be investigated using a cardiotoxin based injury model of skeletal muscle. To study the function of one gene during the regeneration of skeletal muscle, transgenic mouse models are mostly used. Here, we present an alternative method to transgenic mice, to investigate the gene function in satellite cells during regeneration, e.g., in cases where transgenic mice are not available. We combine the cardiotoxin mediated injury of a specific skeletal muscle with the injection of a self-delivering siRNA into the regenerating muscle which is then taken up by satellite cells among other cells. Thereby, we provide a method to analyze gene function in satellite cells during regeneration under physiological conditions without the need for transgenic mice.

A typical result of a hematoxylin and eosin (H&amp;E) staining of a tibialis anterior muscle before and after cardiotoxin mediated injury is presented in Figure 4A,B. In the control muscles, the architecture of the muscle is intact as seen by the localization of the nuclei in the periphery of the myofibers and the lack of accumulation of mononucleated cells in the interstitial space (Figure 4A). 7 days after cardiotoxin mediated injury new myofibers are formed marked by centrally located nuclei (Figure 4B). Furthermore, an accumulation of mononucleated cells can be observed, consisting mostly of satellite cells but also non-myogenic cells like immune cells. The whole muscle should be injured characterized by the central location of all myonuclei and the accumulation of mononucleated cells on the whole muscle section.
To characterize the progression and success of the regeneration process, several immunofluorescent staining using markers of regeneration can be performed. The number of satellite cells can be evaluated by staining for Pax7, the canonical marker for satellite cells (Figure 4C,D). Three days after injury the number of satellite cells increases (Figure 4D), satellite cells are not located under the basal lamina anymore. To further analyze the regeneration process, newly formed myofibers can be stained with antibodies directed to developmental myosin (Figure 4E). Newly formed myofibers display centrally located nuclei and a strong expression of developmental myosin. As the myofibers mature, expression of developmental myosin decreases, the diameter of the myofiber increases while nuclei are migrating to the periphery of the myofibers.
The influence of a single gene on the process of regeneration can be investigated using transgenic mice or as shown here by injection of a self-delivering siRNA. Fluorescently labeled self-delivering scrambled control siRNA was injected into the regenerating tibialis anterior muscle at day 3 after injury, the time point when the proliferation of satellite cells peaks. Two days after injection of the siRNA satellite cells were analyzed for the presence of the fluorescently labeled siRNA (Figure 4F). We analyzed how many satellite cells had taken up the fluorescently labeled siRNA two days after injection into the regenerating muscle (Figure 5). About 75% of all satellite cells in the regenerating muscle were positive for the fluorescently labeled siRNA. Furthermore, we determined that about 74% of all regenerating myofibers were positive for the fluorescently labeled siRNA suggesting that either 74% of the regenerating myofibers had taken up the siRNA or that siRNA-positive satellite cells either had fused with each other to form new myofibers or that fusion with already existing regenerating myofibers occurred (Figure 5). This suggests that satellite cells take up the self-delivering siRNA and that the siRNA persists in the satellite cells for at least two days.
<img alt="Animal experiment setup: rodent under anesthesia, ultrasound and syringe used for injection, research." class="xfigimg" src="/files/ftp_upload/60194/60194fig01.jpg" data-altupdatedat="1747911167000" data-alt="Figure 1"> 
Figure 1: Injection of cardiotoxin into the tibialis anterior muscle. (A) Mice are anesthetized by inhalation of isoflurane and the lower limb is shaved. (B) A 29 G needle is injected into the tibialis anterior muscle (C) and moved along the tibia bone during injection of the cardiotoxin solution. <a href="https://www.jove.com/files/ftp_upload/60194/60194fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Mouse tissue dissection sequence; steps in biological sample preparation and analysis." class="xfigimg" src="/files/ftp_upload/60194/60194fig02.jpg" data-altupdatedat="1747911167000" data-alt="Figure 2"> 
Figure 2: Steps involved in dissection and freezing of the injured tibialis anterior muscle. (A) The hind limb muscles are exposed (B) and the fascia surrounding the tibialis anterior muscle is ripped to expose the muscle. (C) After removing the muscle, it is cut at the mid belly region into two halves of similar size using sharp straight scissors (D). (E) The halves of the dissected muscle are embedded in an aluminum foil mold filled with freezing solution and frozen in liquid nitrogen (F). <a href="https://www.jove.com/files/ftp_upload/60194/60194fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Microtomy process: cryostat setup, sectioning blade, and tissue sample on slide in histological analysis." class="xfigimg" src="/files/ftp_upload/60194/60194fig03.jpg" data-altupdatedat="1747911167000" data-alt="Figure 3"> 
Figure 3: Equipment and steps involved in cryosectioning of tibialis anterior muscles. (A) The chamber temperature of the cryostat is set to -21 °C. (B) The tibialis anterior muscle in the freezing solution is mounted on the sample holder. (C) Sections of 14 µm thickness are attached to glass microscope slides. <a href="https://www.jove.com/files/ftp_upload/60194/60194fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Muscle tissue histology and immunofluorescence, comparing healthy vs. diseased cellular morphology." class="xfigimg" src="/files/ftp_upload/60194/60194fig04.jpg" data-altupdatedat="1747911167000" data-alt="Figure 4"> 
Figure 4: Representative images of regenerating tibialis anterior muscles. H&amp;E staining of tibialis anterior muscles before (A) and 7 days after cardiotoxin injury (B). (C) In uninjured muscle Pax7 positive (in red) satellite cells are located underneath the basal lamina marked by laminin (in green). Nuclei are counterstained with DAPI (in blue). Pax7 positive cells are marked by an arrow. (D) After 10 days of regeneration, myofibers are characterized by centrally located nuclei (in blue), Pax7 is shown in red, laminin in green. Pax7 positive cells are marked by an arrow. (E) Newly formed myofibers express developmental myosin (in red), laminin in green, nuclei are counterstained with DAPI (in blue). (F) Fluorescently labeled self-delivering siRNA (siRED, depicted in red) is still found in satellite cells (Pax7 positive, in green, marked by an arrow in the inset) 2 days after injection of the self-delivering siRNA into regenerating tibialis anterior muscle (injection at day 3 after cardiotoxin mediated injury). Laminin is shown in white, nuclei are counterstained with DAPI (in blue). Scale bar = 100 µm (A and B), 50 µm (C-E), 25 µm (F). <a href="https://www.jove.com/files/ftp_upload/60194/60194fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Bar chart and table showing siRNA positive cell percentages in Pax7+ cells and regenerating myofibers." class="xfigimg" src="/files/ftp_upload/60194/60194fig05.jpg" data-altupdatedat="1747911167000" data-alt="Figure 5"> 
Figure 5: Quantification of uptake of fluorescently labeled siRNA. (A) Quantification of siRNA uptake by satellite cells (Pax7+ cells) and regenerating myofibers 2 days after injection into regenerating skeletal muscle. Injection was performed at day 3 after cardiotoxin mediated injury. n = 3, error bars show SEM. (B) Quantification underlying the graph depicted in (A). <a href="https://www.jove.com/files/ftp_upload/60194/60194fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Analyzing Satellite Cell Function During Skeletal Muscle Regeneration by Cardiotoxin Injury and Injection of Self-delivering siRNA In Vivo at JoVE.com

Analyzing Satellite Cell Function During Skeletal Muscle Regeneration by Cardiotoxin Injury and Injection of Self-delivering siRNA In Vivo

We describe an in vivo method to investigate the functional loss of a specific gene in satellite cells by using a combination of cardiotoxin mediated injury of skeletal muscle and the injection of a self-delivering siRNA.

Skeletal muscle possesses an enormous capacity to regenerate after injury. This process is mainly driven by muscle stem cells, also termed satellite ...

analyzing-satellite-cell-function-during-skeletal-muscle-regeneration

Research

JoVE Journal

Genetics

14.8K Views.  Leibniz-Institute on Aging - Fritz-Lipmann-Institute. Deze techniek maakt de combinatie van de intramusculaire injectie van het slangengif cardiotoxine, de intramusculaire injectie van zelfleverende siRNA, waardoor de analyse van regeneratie van skeletspieren mogelijk is. De toepassing van zelfleverend siRNA is een elegante manier om het functionele verlies van enkele genen tijdens spierregeneratie te onderzoeken, waardoor transgene dieren worden vermeden. Na bevestiging van een gebrek aan respons op teensnuifje, scheert u het injectiegebied van...

Video: Analyzing Satellite Cell Function During Skeletal Muscle Regeneration by Cardiotoxin Injury and Injection of Self-delivering siRNA In Vivo

Isolation and Characterization of Satellite Cells from Rat Head Branchiomeric Muscles

Fibrosis and defective muscle regeneration can hamper the functional recovery of the soft palate muscles after cleft palate repair. This causes persistent problems in speech, swallowing, and sucking. In vitro culture systems that allow the study of satellite cells (myogenic stem cells) from head muscles are crucial to develop new therapies based on tissue engineering to promote muscle regeneration after surgery. These systems will offer new perspectives for the treatment of cleft palate patients. A protocol for the isolation, culture and differentiation of satellite cells from head muscles is presented. The isolation is based on enzymatic digestion and trituration to release the satellite cells. In addition, this protocol comprises an innovative method using extracellular matrix gel coatings of millimeter size, which requires only low numbers of satellite cells for differentiation assays.

This protocol describes the isolation of satellite cells from branchiomeric head muscles of a 9 week-old rat. The muscles originate from different branchial arches. Subsequently, the satellite cells are cultured on a spot coating of millimeter size to study their differentiation. This approach avoids the expansion and passaging of satellite cells.

Fibrosis and defective muscle regeneration can hamper the functional recovery of the soft palate muscles after cleft palate repair. This causes persistent ...

Developmental Biology

Induction of Acute Skeletal Muscle Regeneration by Cardiotoxin Injection

Skeletal muscle regeneration is a physiological process that occurs in adult skeletal muscles in response to injury or disease. Acute injury-induced skeletal muscle regeneration is a widely used, powerful model system to study the events involved in muscle regeneration as well as the mechanisms and different players. Indeed, a detailed knowledge of this process is essential for a better understanding of the pathological conditions that lead to skeletal muscle degeneration, and it aids in identifying new targeted therapeutic strategies. The present work describes a detailed and reproducible protocol to induce acute skeletal muscle regeneration in mice through a single intramuscular injection of cardiotoxin (CTX). CTX belongs to the family of snake venom toxins and causes myolysis of myofibers, which eventually triggers the regeneration events. The dynamics of skeletal muscle regeneration is evaluated by histological analysis of muscle sections. The protocol also illustrates the experimental procedures for dissecting, freezing, and cutting the Tibialis Anterior muscle, as well as the routine Hematoxylin &#38; Eosin staining that is widely used for subsequent morphological and morphometric analysis.

This manuscript describes a detailed protocol to induce acute skeletal muscle regeneration in adult mice and subsequent manipulations of the muscles, such as dissection, freezing, cutting, routine staining, and myofiber cross-sectional area analysis.

Skeletal muscle regeneration is a physiological process that occurs in adult skeletal muscles in response to injury or disease. Acute injury-induced ...

Transplantation of Induced Pluripotent Stem Cell-derived Mesoangioblast-like Myogenic Progenitors in Mouse Models of Muscle Regeneration

Patient-derived iPSCs could be an invaluable source of cells for future autologous cell therapy protocols. iPSC-derived myogenic stem/progenitor cells similar to pericyte-derived mesoangioblasts (iPSC-derived mesoangioblast-like stem/progenitor cells: IDEMs) can be established from iPSCs generated from patients affected by different forms of muscular dystrophy. Patient-specific IDEMs can be genetically corrected with different strategies (e.g. lentiviral vectors, human artificial chromosomes) and enhanced in their myogenic differentiation potential upon overexpression of the myogenesis regulator MyoD. This myogenic potential is then assessed in vitro with specific differentiation assays and analyzed by immunofluorescence. The regenerative potential of IDEMs is further evaluated in vivo, upon intramuscular and intra-arterial transplantation in two representative mouse models displaying acute and chronic muscle regeneration. The contribution of IDEMs to the host skeletal muscle is then confirmed by different functional tests in transplanted mice. In particular, the amelioration of the motor capacity of the animals is studied with treadmill tests. Cell engraftment and differentiation are then assessed by a number of histological and immunofluorescence assays on transplanted muscles. Overall, this paper describes the assays and tools currently utilized to evaluate the differentiation capacity of IDEMs, focusing on the transplantation methods and subsequent outcome measures to analyze the efficacy of cell transplantation.

Induced pluripotent stem cell (iPSC)-derived myogenic progenitors are promising candidates for cell therapy strategies to treat muscular dystrophies. This protocol describes transplantation and functional measurements required to evaluate the engraftment and differentiation of iPSC-derived mesoangioblasts (a type of muscle progenitors) in mouse models of acute and chronic muscle regeneration.

Patient-derived iPSCs could be an invaluable source of cells for future autologous cell therapy protocols. iPSC-derived myogenic stem/progenitor cells ...

Bioengineering

Intramyocardial Cell Delivery: Observations in Murine Hearts

Previous studies showed that cell delivery promotes cardiac function amelioration by release of cytokines and factors that increase cardiac tissue revascularization and cell survival. In addition, further observations revealed that specific stem cells, such as cardiac stem cells, mesenchymal stem cells and cardiospheres have the ability to integrate within the surrounding myocardium by differentiating into cardiomyocytes, smooth muscle cells and endothelial cells.
Here, we present the materials and methods to reliably deliver noncontractile cells into the left ventricular wall of immunodepleted mice. The salient steps of this microsurgical procedure involve anesthesia and analgesia injection, intratracheal intubation, incision to open the chest and expose the heart and delivery of cells by a sterile 30-gauge needle and a precision microliter syringe.
Tissue processing consisting of heart harvesting, embedding, sectioning and histological staining showed that intramyocardial cell injection produced a small damage in the epicardial area, as well as in the ventricular wall. Noncontractile cells were retained into the myocardial wall of immunocompromised mice and were surrounded by a layer of fibrotic tissue, likely to protect from cardiac pressure and mechanical load.

Intramyocardial cell delivery in murine models of cardiovascular diseases, such as hypertension or myocardial infarction, is widely used to test the therapeutic potential of different cell types in regenerative studies. Therefore, a detailed description and a clear visualization of this surgical procedure will help to define the limits and advantages of cardiovascular cell therapeutic analyses in small rodents.

Previous studies showed that cell delivery promotes cardiac function amelioration by release of cytokines and factors that increase cardiac tissue ...

Medicine

Applications of In Vivo Functional Testing of the Rat Tibialis Anterior for Evaluating Tissue Engineered Skeletal Muscle Repair

Despite the regenerative capacity of skeletal muscle, permanent functional and/or cosmetic deficits (e.g., volumetric muscle loss (VML) resulting from traumatic injury, disease and various congenital, genetic and acquired conditions are quite common. Tissue engineering and regenerative medicine technologies have enormous potential to provide a therapeutic solution. However, utilization of biologically relevant animal models in combination with longitudinal assessments of pertinent functional measures are critical to the development of improved regenerative therapeutics for treatment of VML-like injuries. In that regard, a commercial muscle lever system can be used to measure length, tension, force and velocity parameters in skeletal muscle. We used this system, in conjunction with a high power, bi-phase stimulator, to measure in vivo force production in response to activation of the anterior crural compartment of the rat hindlimb. We have previously used this equipment to assess the functional impact of VML injury on the tibialis anterior (TA) muscle, as well as the extent of functional recovery following treatment of the injured TA muscle with our tissue engineered muscle repair (TEMR) technology. For such studies, the left foot of an anaesthetized rat is securely anchored to a footplate linked to a servomotor, and the common peroneal nerve is stimulated by two percutaneous needle electrodes to elicit muscle contraction and dorsiflexion of the foot. The peroneal nerve stimulation-induced muscle contraction is measured over a range of stimulation frequencies (1-200 Hz), to ensure an eventual plateau in force production that allows for an accurate determination of peak tetanic force. In addition to evaluation of the extent of VML injury as well as the degree of functional recovery following treatment, this methodology can be easily applied to study diverse aspects of muscle physiology and pathophysiology. Such an approach should assist with the more rational development of improved therapeutics for muscle repair and regeneration.

Applications of In Vivo Functional Testing of the Rat Tibialis Anterior for Evaluating Tissue Engineered Skeletal Muscle Repair

We describe an in vivo protocol to measure dorsiflexion of the foot following stimulation of the peroneal nerve and contraction of the anterior crural compartment of the rat hindlimb. Such measurements are an indispensable translational tool for evaluating skeletal muscle pathology and tissue engineering approaches to muscle repair and regeneration.

Despite the regenerative capacity of skeletal muscle, permanent functional and/or cosmetic deficits (e.g., volumetric muscle loss (VML) resulting from ...

Minimally Invasive Muscle Embedding (MIME) - A Novel Experimental Technique to Facilitate Donor-Cell-Mediated Myogenesis

Skeletal muscle possesses regenerative capacity due to tissue-resident, muscle-fiber-generating (myogenic) satellite cells (SCs), which can form new muscle fibers under the right conditions. Although SCs can be harvested from muscle tissue and cultured in vitro, the resulting myoblast cells are not very effective in promoting myogenesis when transplanted into host muscle. Surgically exposing the host muscle and grafting segments of donor muscle tissue, or the isolated muscle fibers with their SCs onto host muscle, promotes better myogenesis compared to myoblast transplantation. We have developed a novel technique that we call Minimally Invasive Muscle Embedding (MIME). MIME involves passing a surgical needle through the host muscle, drawing a piece of donor muscle tissue through the needle track, and then leaving the donor tissue embedded in the host muscle so that it may act as a source of SCs for the host muscle. Here we describe in detail the steps involved in performing MIME in an immunodeficient mouse model that expresses a green fluorescent protein (GFP) in all of its cells. Immunodeficiency in the host mouse reduces the risk of immune rejection of the donor tissue, and GFP expression enables easy identification of the host muscle fibers (GFP+) and donor-cell-derived muscle fibers (GFP-). Our pilot data suggest that MIME can be used to implant an extensor digitorum longus (EDL) muscle from a donor mouse into the tibialis anterior (TA) muscle of a host mouse. Our data also suggest that when a myotoxin (barium chloride, BaCl2) is injected into the host muscle after MIME, there is evidence of donor-cell-derived myogenesis in the host muscle, with approximately 5%, 26%, 26% and 43% of the fibers in a single host TA muscle showing no host contribution, minimal host contribution, moderate host contribution, and maximal host contribution, respectively.

Minimally Invasive Muscle Embedding MIME - A Novel Experimental Technique to Facilitate Donor-Cell-Mediated Myogenesis

We describe a novel experimental technique that we call Minimally Invasive Muscle Embedding (MIME), which is based on the evidence that skeletal muscle tissue contains viable myogenic cells that can facilitate donor-cell-mediated myogenesis when implanted into a host muscle.

Skeletal muscle possesses regenerative capacity due to tissue-resident, muscle-fiber-generating (myogenic) satellite cells (SCs), which can form new ...

Isolation and Culture of Individual Myofibers and their Satellite Cells from Adult Skeletal Muscle

Muscle regeneration in the adult is performed by resident stem cells called satellite cells. Satellite cells are defined by their position between the basal lamina and the sarcolemma of each myofiber. Current knowledge of their behavior heavily relies on the use of the single myofiber isolation protocol. In 1985, Bischoff described a protocol to isolate single live fibers from the Flexor Digitorum Brevis (FDB) of adult rats with the goal to create an in vitro system in which the physical association between the myofiber and its stem cells is preserved 1. In 1995, Rosenblattmodified the Bischoff protocol such that myofibers are singly picked and handled separately after collagenase digestion instead of being isolated by gravity sedimentation 2, 3. The Rosenblatt or Bischoff protocol has since been adapted to different muscles, age or conditions 3-6. The single myofiber isolation technique is an indispensable tool due its unique advantages. First, in the single myofiber protocol, satellite cells are maintained beneath the basal lamina. This is a unique feature of the protocol as other techniques such as Fluorescence Activated Cell Sorting require chemical and mechanical tissue dissociation 7. Although the myofiber culture system cannot substitute for in vivo studies, it does offer an excellent platform to address relevant biological properties of muscle stem cells. Single myofibers can be cultured in standard plating conditions or in floating conditions. Satellite cells on floating myofibers are subjected to virtually no other influence than the myofiber environment. Substrate stiffness and coating have been shown to influence satellite cells' ability to regenerate muscles 8, 9 so being able to control each of these factors independently allows discrimination between niche-dependent and -independent responses. Different concentrations of serum have also been shown to have an effect on the transition from quiescence to activation. To preserve the quiescence state of its associated satellite cells, fibers should be kept in low serum medium 1-3. This is particularly useful when studying genes involved in the quiescence state. In serum rich medium, satellite cells quickly activate, proliferate, migrate and differentiate, thus mimicking the in vivo regenerative process 1-3. The system can be used to perform a variety of assays such as the testing of chemical inhibitors; ectopic expression of genes by virus delivery; oligonucleotide based gene knock-down or live imaging. This video article describes the protocol currently used in our laboratory to isolate single myofibers from the Extensor Digitorum Longus (EDL) muscle of adult mice (6-8 weeks old).

Isolation and culture of myofibers is the gold standard in vitro system to study the transition of satellite cells through quiescence, activation and differentiation. Importantly, the single myofiber culture system preserves the myofiber/stem cell association, which is an essential component of the muscle stem cell niche.

Muscle  regeneration in the adult is performed by resident stem cells called satellite  cells. Satellite cells are defined by  their position between the ...

Biology

Isolation, Culture, and Transplantation of Muscle Satellite Cells

Muscle satellite cells are a stem cell population required for postnatal skeletal muscle development and regeneration, accounting for 2-5% of sublaminal nuclei in muscle fibers. In adult muscle, satellite cells are normally mitotically quiescent. Following injury, however, satellite cells initiate cellular proliferation to produce myoblasts, their progenies, to mediate the regeneration of muscle. Transplantation of satellite cell-derived myoblasts has been widely studied as a possible therapy for several regenerative diseases including muscular dystrophy, heart failure, and urological dysfunction. Myoblast transplantation into dystrophic skeletal muscle, infarcted heart, and dysfunctioning urinary ducts has shown that engrafted myoblasts can differentiate into muscle fibers in the host tissues and display partial functional improvement in these diseases. Therefore, the development of efficient purification methods of quiescent satellite cells from skeletal muscle, as well as the establishment of satellite cell-derived myoblast cultures and transplantation methods for myoblasts, are essential for understanding the molecular mechanisms behind satellite cell self-renewal, activation, and differentiation. Additionally, the development of cell-based therapies for muscular dystrophy and other regenerative diseases are also dependent upon these factors.

However, current prospective purification methods of quiescent satellite cells require the use of expensive fluorescence-activated cell sorting (FACS) machines. Here, we present a new method for the rapid, economical, and reliable purification of quiescent satellite cells from adult mouse skeletal muscle by enzymatic dissociation followed by magnetic-activated cell sorting (MACS). Following isolation of pure quiescent satellite cells, these cells can be cultured to obtain large numbers of myoblasts after several passages. These freshly isolated quiescent satellite cells or ex vivo expanded myoblasts can be transplanted into cardiotoxin (CTX)-induced regenerating mouse skeletal muscle to examine the contribution of donor-derived cells to regenerating muscle fibers, as well as to satellite cell compartments for the examination of self-renewal activities.

The isolation and culture of a pure population of quiescent satellite cells, a muscle stem cell population, is essential to the understanding of muscle stem cell biology and regeneration, as well as stem cell transplantation for therapies in muscular dystrophy and other degenerative diseases.&#160;

Muscle satellite cells are a stem cell population required for postnatal skeletal muscle development and regeneration, accounting for 2-5% of sublaminal ...

In Situ Immunofluorescent Staining of Autophagy in Muscle Stem Cells

Increasing evidence points to autophagy as a crucial regulatory process to preserve tissue homeostasis. It is known that autophagy is involved in skeletal muscle development and regeneration, and the autophagic process has been described in several muscular pathologies and age-related muscle disorders. A recently described block of the autophagic process that correlates with the functional exhaustion of satellite cells during muscle repair supports the notion that active autophagy is coupled with productive muscle regeneration. These data uncover the crucial role of autophagy in satellite cell activation during muscle regeneration in both normal and pathological conditions, such as muscular dystrophies. Here, we provide a protocol to monitor the autophagic process in the adult Muscle Stem Cell (MuSC) compartment during muscle regenerative conditions. This protocol describes the setup methodology to perform in situ immunofluorescence imaging of LC3, an autophagy marker, and MyoD, a myogenic lineage marker, in muscle tissue sections from control and injured mice. The methodology reported allows for monitoring the autophagic process in one specific cell compartment, the MuSC compartment, which plays a central role in orchestrating muscle regeneration.

In Situ Immunofluorescent Staining of Autophagy in Muscle Stem Cells

Active autophagy is associated with productive muscle regeneration, which is essential for Muscle Stem Cell (MuSC) activation. Here, we provide a protocol for the in situ detection of LC3, an autophagy marker in MyoD-positive MuSCs of muscle tissue sections from control and injured mice.

Increasing evidence points to autophagy as a crucial regulatory process to preserve tissue homeostasis. It is known that autophagy is involved in skeletal ...

Single-Cell CyTOF Analysis of Myogenic Progenitors in Muscle Regeneration

Identification and Analysis of Myogenic Progenitors In Vivo During Acute Skeletal Muscle Injury by High-Dimensional Single-Cell Mass Cytometry

Skeletal muscle regeneration is a dynamic process driven by adult muscle stem cells and their progeny. Mostly quiescent at a steady state, adult muscle stem cells become activated upon muscle injury. Following activation, they proliferate, and most of their progeny differentiate to generate fusion-competent muscle cells while the remaining self-renews to replenish the stem cell pool. While the identity of muscle stem cells was defined more than a decade ago, based on the co-expression of cell surface markers, myogenic progenitors were identified only recently using high-dimensional single-cell approaches. Here, we present a single-cell mass cytometry (cytometry by time of flight [CyTOF]) method to analyze stem cells and progenitor cells in acute muscle injury to resolve the cellular and molecular dynamics that unfold during muscle regeneration. This approach is based on the simultaneous detection of novel cell surface markers and key myogenic transcription factors whose dynamic expression enables the identification of activated stem cells and progenitor cell populations that represent landmarks of myogenesis. Importantly, a sorting strategy based on detecting cell surface markers CD9 and CD104 is described, enabling prospective isolation of muscle stem and progenitor cells using fluorescence-activated cell sorting (FACS) for in-depth studies of their function. Muscle progenitor cells provide a critical missing link to study the control of muscle stem cell fate, identify novel therapeutic targets for muscle diseases, and develop cell therapy applications for regenerative medicine. The approach presented here can be applied to study muscle stem and progenitor cells in vivo in response to perturbations, such as pharmacological interventions targeting specific signaling pathways. It can also be used to investigate the dynamics of muscle stem and progenitor cells in animal models of muscle diseases, advancing our understanding of stem cell diseases and accelerating the development of therapies.

Author Spotlight: Investigating Cellular and Molecular Dynamics During Muscle Regeneration Using Cutting-Edge Single-Cell Technologies

The protocol presented here enables the identification and high-dimensional analysis of muscle stem and progenitor cells by single-cell mass cytometry and their purification by FACS for in-depth studies of their function. This approach can be applied to study regeneration dynamics in disease models and test the efficacy of pharmacological interventions.

Skeletal muscle regeneration is a dynamic process driven by adult muscle stem cells and their progeny. Mostly quiescent at a steady state, adult muscle ...

Analyzing Satellite Cell Function During Skeletal Muscle Regeneration by Cardiotoxin Injury and Injection of Self-delivering siRNA In Vivo

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